a lack of parasitic reduction in the obligate parasitic ...a lack of parasitic reduction in the...

A Lack of Parasitic Reduction in the Obligate ParasiticGreen Alga HelicosporidiumJean-Francois Pombert1¤, Nicolas Achille Blouin2, Chris Lane2, Drion Boucias3, Patrick J. Keeling1*

1 Canadian Institute for Advanced Research, Department of Botany, University of British Columbia, Vancouver, British Columbia, Canada, 2 Department of Biological

Sciences, University of Rhode Island, Kingston, Rhode Island, United States of America, 3 Entomology and Nematology Department, University of Florida, Gainesville,

Florida, United States of America

Abstract

The evolution of an obligate parasitic lifestyle is often associated with genomic reduction, in particular with the loss offunctions associated with increasing host-dependence. This is evident in many parasites, but perhaps the most extremetransitions are from free-living autotrophic algae to obligate parasites. The best-known examples of this are theapicomplexans such as Plasmodium, which evolved from algae with red secondary plastids. However, an analogoustransition also took place independently in the Helicosporidia, where an obligate parasite of animals with an intracellularinfection mechanism evolved from algae with green primary plastids. We characterised the nuclear genome ofHelicosporidium to compare its transition to parasitism with that of apicomplexans. The Helicosporidium genome is smalland compact, even by comparison with the relatively small genomes of the closely related green algae Chlorella andCoccomyxa, but at the functional level we find almost no evidence for reduction. Nearly all ancestral metabolic functions areretained, with the single major exception of photosynthesis, and even here reduction is not complete. The great majority ofgenes for light-harvesting complexes, photosystems, and pigment biosynthesis have been lost, but those for otherphotosynthesis-related functions, such as Calvin cycle, are retained. Rather than loss of whole function categories, thepredominant reductive force in the Helicosporidium genome is a contraction of gene family complexity, but even here mostlosses affect families associated with genome maintenance and expression, not functions associated with host-dependence.Other gene families appear to have expanded in response to parasitism, in particular chitinases, including those predictedto digest the chitinous barriers of the insect host or remodel the cell wall of Helicosporidium. Overall, the Helicosporidiumgenome presents a fascinating picture of the early stages of a transition from free-living autotroph to parasitic heterotrophwhere host-independence has been unexpectedly preserved.

Citation: Pombert J-F, Blouin NA, Lane C, Boucias D, Keeling PJ (2014) A Lack of Parasitic Reduction in the Obligate Parasitic Green Alga Helicosporidium. PLoSGenet 10(5): e1004355. doi:10.1371/journal.pgen.1004355

Editor: Joseph Heitman, Duke University Medical Center, United States of America

Received January 2, 2014; Accepted March 21, 2014; Published May 8, 2014

Copyright: � 2014 Pombert et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permitsunrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This work was supported by a grant from the Canadian Institutes of Health Research to PJK (MOP-42517; http://www.cihr-irsc.gc.ca/). The funders hadno role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing Interests: The authors have declared that no competing interests exist.

* E-mail: [email protected]

¤ Current address: Department of Biological and Chemical Sciences, Illinois Institute of Technology, Chicago, Illinois, United States of America

Introduction

Helicosporidia are parasitic protists characterized by mature

discoid cysts each containing a single filamentous and three

ovoid cells [1,2]. These parasites invade their invertebrate

hosts per os and initiate their replicative stage within the

digestive tract [3,4]. The cysts, triggered by chemical changes

in the gut, dehisce and release both the ovoid cells and filament

cell. The ovoid cells remain in the gut lumen whereas the

uncoiled and barbed filamentous cells penetrate the peri-

trophic membrane and become anchored to the host midgut

cells. Over time the filamentous cells migrate through the

midgut epithelium, breach the basement membrane, and

invade the hemocoel. In the hemocoel the filament cells will

transition to a vegetative stage that replicates by autosporula-

tion; a select number at the four-cell stage will differentiate to

the infectious cyst stage characterized by the three ovoid and a

single filament cell [5,6]. Unlike many parasites the vegetative

cells of Helicosporidia can be cultured readily on defined

media with limited nutrients, suggesting that despite being a

parasitic species, they have retained a diverse slate of

metabolic pathways allowing for saprobic growth.

The evolutionary origin of Helicosporidia remained uncertain

for nearly 100 years since their initial description, although various

characters were used to suggest some relationship with microspo-

ridian, sporozoan, and myxosporidian parasites. Recently, how-

ever, ultrastructural observations surprisingly revealed that the

vegetative state of Helicosporidium cells is similar to that of the

achlorophylous trebouxiophyte green alga Prototheca [1], and

subsequent phylogenetic inferences derived from actin/tubulin

and plastid sequences strongly confirmed this affiliation [7,8]. The

discovery that Helicosporidia are trebouxiophycean green algae

raises some interesting questions about the evolution of parasitism:

within this single lineage are found free-living autotrophs like most

other green algae, but also a variety of symbiotic species,

opportunistic pathogens, and perhaps even obligate intracellular

parasites, all of which diversified within a relatively narrow

evolutionary timescale. The transformation from free-living to

parasitic lifestyles often includes the shedding of metabolic

functions that are no longer required as the parasite relies

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increasingly on its host for energy and nutrients [9]. The parasitic

relationship may be opportunistic at first, but can switch to being

obligate upon reaching a certain threshold of host-dependence,

after which the formerly free-living organism can no longer revert

to its previous lifestyle due to the ratchet-like nature of these losses.

We do not often think of photosynthetic organisms with

progenitors for parasitic ones, but a variety or parasitic lineages

had at one time photosynthetic ancestors, including oomycetes,

several dinoflagellates, and most famously the apicomplexan

parasites such as the malaria parasite, Plasmodium (see [10,11]

and references therein). One of the first things to be lost in

photosynthetic species is presumably their ability to harvest energy

from light and fix carbon. Harnessing light from within large-

bodied hosts is probably very difficult if not impossible, and the

resulting metabolic deficit must lead to a significant shift in the

balance between the host and parasite. Some of these lineages (e.g.

oomycetes) probably evolved through a heterotrophic intermedi-

ate, but others possibly began their association with animals as

phototrophs. How the transformation to parasites took place is of

great interest, but unfortunately because it happened so long ago

(around 1 bya for Plasmodium [12]) and is now so complete, the

critical early stages have long been wiped away. Helicosporidia, in

contrast, appear to have evolved from free-living autotrophs

relatively recently [13,14], and might therefore provide some

interesting insights. Fossils records and molecular clock analyses

suggest that Trebouxiophytes as a group arose in the early

Neoproterozoic [13], from which the trebouxiophycean subgroup

Chorellales later emerged around 100 million years ago (mya)

[14]. Both Helicosporidium and the non-photosynthetic trebouxio-

phycean Prototheca arose from within the Chorellales [13], so the

adaption to parasitism in Helicosporidia occurred less than

100 mya.

To specifically investigate how the metabolic and proteomic

complexity of pathogenic Helicosporidia are distinguished from

their free-living and symbiotic trebouxiophycean relatives, we

sequenced the genome and transcriptome of Helicosporidium sp.

ATCC50920, a parasite of the black fly Simulium jonesi [1,15]. We

show that the Helicosporidium genome is 2.5-fold smaller than

genomes from the free-living and symbiotic trebouxiophytes,

Coccomyxa subellipsoidea C-169 [16] and Chlorella variabilis NC64A

[17], which are themselves extremely small for trebouxiophyte

genomes. However, the reduction of the Helicosporidium genome is

not tied to a massive reduction in metabolic functions: despite its

small genome size and parasitic nature, it surprisingly still encodes

all major metabolic pathways, with the exception of a small

number specifically related to photosynthesis. Even here, the

reduction is not complete: all genes relating to light harvesting and

electron transport are missing, but the Helicosporidium carbon

fixation pathway is nearly complete but for the lack of ribulose-1,

5-bisphosphate carboxylase/oxygenase (RuBisCO) and a pyruvate

kinase. The smaller size of the Helicosporidium genome can be

attributed to a greater degree of genome compaction (e.g. fewer

and smaller introns, and smaller intergenic regions), and most

significantly to a lower complexity of gene families, particularly

those related to DNA packaging/replication pathways. We also

show that the gene family complexity of other metabolic pathways

has increased, in particular relating to chitin metabolism, which

likely represented a key development in the ability of Helicospor-

idium to develop in the insect host. Overall, these results give our

first view into the early stage in the transition from a free-living

autotroph to an obligate pathogen.

Results

General features of the Helicosporidium draft genomeShotgun Illumina reads of total DNA were assembled into

11,717 contigs totalling 13,684,556 bp (62.2% GC). Contamina-

tion filters suggest a maximum of 1% overall contamination,

located in the smaller-sized contigs. Removal of the mitochondrial

and plastid genomes and filtering of the small contigs resulted in

5,666 contigs of at least 500 bp in size (12,373,820 bp total; N50

3,036 bp, 61.7% GC), with an average coverage of 626 (Table 1).

Based on the current data, we estimate the Helicosporidium genome

at a maximum size of 1760.5 Mbp. This corresponds well to a

genome size estimate of 13 Mbp, derived from karyotype

visualisation by clamped homogeneous electrical field (CHEF)

electrophoresis [18]. A total of 6,035 protein-encoding genes were

predicted among the assembled 12.4 Mbp, with an average of 2.3

exons (366 bp/exon) and 1.3 introns (168 bp/intron) per gene.

Coding density in the Helicosporidium genome is high (0.487 gene/

kb) compared to the free-living and symbiotic trebouxiophytes

Coccomyxa (0.197 gene/kb) and Chlorella (0.212 gene/kb), but lower

than that of the 12 Mbp genomes of the picoplanktonic

prasinophycean green algae in the genus Ostreococcus (0.626 and

0.580 gene/kb; Table 1, Figure 1). Identifiable transposable

elements are rare in the assembled Helicosporidium contigs, although

micro- and minisatellites and regions of generally low complexity

were found (Data S1). Comparing the genomic assemblies with the

transcriptome revealed a total of 95.4% of the genes attributed to

known metabolic pathways (Table S1) were found in both datasets,

and only 3.6% were found exclusively in the transcriptome. This

correlated well with the overall percentages of transcriptomic

contigs mapping on the genomic ones ($1000 bp; 92.3%, $

1500 bp; 95.9%), suggesting that the total coding potential of

Helicosporidium is well-represented in the draft genome. The

Helicosporidium genome shares little gene order conservation with

the other green algal genomes: only 30% of the genes located

within its ten largest contigs are arrayed in syntenic clusters with

those of Chlorella (Figure 2), with no apparent metabolic

relationship between the genes present in these clusters.

A surprisingly near-complete metabolic profileThe Helicosporidium genome is small: it is approximately 2.5

times smaller than the two other complete trebouxiophyte

genomes, Coccomyxa and Chlorella (Table 1, Figure 1), which are

themselves at the extremely low end of the spectrum of estimated

genome sizes in this lineage (Figure S1). But this small size is not a

reflection of a severe reduction in metabolic potential. Indeed, the

Author Summary

Helicosporidium is a highly-adapted obligate parasite ofanimals. Its evolutionary origins were unclear for almost acentury, but molecular analysis ultimately and surprisinglyshowed that it is a green alga, which means it hasundergone an evolutionary transition from autotrophy toparasitism comparable to that of the malaria parasitePlasmodium and its relatives. Such transitions are oftenassociated with the loss of biological functions that are nolonger necessary in their novel environment and with thedevelopment of molecular mechanisms, sometimes quitesophisticated, to invade and take advantage of their hosts.Yet, very little is actually known about the early stages ofthe transition of a free-living organism to an obligateintracellular parasite. Here we sequenced the genome andtranscriptome of Helicosporidium, and use it to show thatthe outcome of this transition is quite different from thatof Plasmodium.

The Obligate Parasitic Green Alga Helicosporidium


Helicosporidium genome encodes almost all of the major biological

functions that are shared between the genomes of its treboux-

iophycean relatives and that of the chlorophycean green alga

Chlamydomonas reinhardtii (Table S1).

Gene loss in the Helicosporidium genome is significantly concen-

trated in photosynthesis-related pathways, and even here gene loss

is surprisingly sparse given its non-photosynthetic, parasitic nature.

The Helicosporidium genome encodes 56% of the plastid-targeted

proteins predicted by the GreenCut2 database [19] (Figure 3),

whereas both the photosynthetic Coccomyxa and Chlorella encode

96% of these proteins. The overall distribution of these losses in

plastid metabolism is not random, but is concentrated on processes

related to light-harvesting (Figure 4, Data S3, S4, S5, S6, S7, S8,

S9, S10, S11). The heme synthesis branch of the tetrapyrrole

pathway is complete in Helicosporidium, but the branch leading to

the biogenesis of chlorophyll has been lost (Figures 4 and S2, Data

S3, S4, S5, S6, S7, S8, S9, S10, S11). Similarly, Helicosporidium

cannot synthesize carotenoids. It does not encode light-harvesting

antenna proteins and photosystems I and II are completely absent,

which parallels the loss of all photosynthesis-related genes in its

plastid genome [20]. Surprisingly however, the Helicosporidium

genome has retained an almost complete carbon fixation pathway

despite lacking two major components rbcL/rbcS coding respec-

tively for the large and small subunits of the ribulose-1,5-

bisphosphate carboxylase oxygenase (RuBisCO) and ppdK, a

pyruvate orthophosphate dikinase involved in pyruvate intercon-

versions in the C4 pathway (Figure 4). Similarly, Helicosporidium has

retained some proteins involved in electron transport and

components of the F-type ATPase and cytochrome b6f

(Figure 4). Starch and fatty acid metabolic pathways are more

or less intact, as is the terpenoid biosynthesis pathway and its

isoprenoid non-mevalonate MEP/DOXP synthesis branch. The

SUF iron-sulfur cluster biosynthetic pathway is conserved as well,

alongside its ISC/NIF mitochondrial counterpart [21,22]. Not

surprisingly the protein import and export systems are intact.

Outside the plastid, Helicosporidium metabolism shows little signs

of significant reduction. The Helicosporidium genome encodes all

proteins required for the biosynthesis of conventional aminoacyl-

tRNAs, except selenocysteine. Despite its conservation across the

green algae, we found no evidence for the presence of a

selenocysteine synthase in the genomic and transcriptomic

Helicosporidium datasets. The o-phosphoseryl-tRNA(sec) kinase also

required for selenocystenyl-tRNA synthesis is missing too, however

all other enzymes involved in the metabolism of selenocompounds

are present. Helicosporidium appears incapable of endogenous RNA

interference and, like the picoeukaryotes Ostreococcus tauri and

Ostreococcus lucimarinus, lacks the genes coding for the Dicer and

Argonaute proteins. These genes are found in single copies in the

Chlorella and Coccomyxa genomes whereas three paralogous copies

of DC1 and AGO1 are found in the Chlamydomonas genome [23].

In Chlamydomonas, this expanded set has been postulated to mediate

the silencing of its numerous transposable elements [24]. The few

losses observed in the remaining Helicosporidium pathways are either

palliated by bypass enzymes or affect the synthesis or homeostasis

of uncommon metabolites (Table S2).

The increased level of compaction and loss of photosynthetic

genes in the Helicosporidium genome cannot explain its 2.5-fold

reduction in genome size: other significant differences in gene

content exist between the parasite and its free-living relatives.

Given that Helicosporidium, Coccomyxa, and Chlorella were found to

encode almost the same overall functional categories of genes in

common with other green algae (Table S1), one possibility is that

Helicosporidium possess fewer and/or smaller gene families. To

investigate the complexity of gene families, we compared the

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Helicosporidium, Chlorella, and Coccomyxa predicted proteomes via an

evolutionary gene network analysis [25]. A total of 100 connected

components, excluding photosynthesis-related products, were

found to exhibit a lower representation in Helicosporidium compared

with its free-living relatives. These were manually curated into 9

functional categories based on annotation of the three gene sets

(Tables S3 and S4, Figure 5). Interestingly, the functional

categories where Helicosporidium has a reduced gene family

complexity are for the most part not what are broadly defined

as ‘operational’ genes where reduction that might be related to

increased dependence on the host. Instead, the most drastic

reductions in Helicosporidium are gene families in functional groups

that are correlated with the size and complexity of the genome.

The few exceptions are in amino acid and some other metabolic

families, but most of the reduction relates to genes involved in

chromosome packing, transcription, translation, post-translational

Figure 1. Inversed correlation between size and coding density in sequenced green algal genomes. Assembled sizes (Mbp) and codingdensities (gene/kbp) are shown on the left and right Y-axis, respectively.doi:10.1371/journal.pgen.1004355.g001

Figure 2. Conserved gene clusters between the Helicosporidium and Chlorella genomes. Only syntenic clusters from the ten largestHelicosporidium contigs are shown. Genes from Helicosporidium are shown on top; Chlorella genes are shown below. Locus_tag prefixes(Helicosporidium, H632_; Chlorella, CHLNDRAFT_) were omitted for clarity (see Data S2 for PFAM product names). The corresponding contigs(Helicosporidium) or scaffolds (Chlorella) are indicated on the left; in the Chlorella scaffolds, adjacent genes are not always labelled incrementally.Genes that are absent from the other genome are colored in light gray. Genes that have been relocated are shown in dark gray. Partial genes areindicated by double daggers (`). In Helicosporidium the c9p0 gene, indicated by an asterisk, is predicted as a single entity encompassing the Chlorella32522 and 36846 genes.doi:10.1371/journal.pgen.1004355.g002



Figure 3. Distribution of GreenCut2 proteins in Helicosporidium. The pathways indicated on the left are from GreenCut2 [19]. Numbers insidethe white and green bars indicate the total number of genes that are absent or present, respectively, from the corresponding pathway in theHelicosporidium genome. The bars are drawn to scale according to the respective percentage of genes that have been retained (green) or lost (white)in the corresponding pathway.doi:10.1371/journal.pgen.1004355.g003

Figure 4. Localization of Helicosporidium losses amongst the major plastid metabolic pathways. Proteins present and absent inHelicosporidium are shown in blue and orange, respectively. The metabolic pathways shown here have been simplified for visualization; only theproteins present in other green algae were considered and signal peptidases from the protein export pathway perform their functions in the cytosol.The KEGG orthology (ko) identifiers [50,51] are: carbon fixation, ko00710; carotenoids, ko00906; fatty acids, ko01040; ile/leu/val, ko00290; phe/tyr/trp,ko00400; protein export, ko03060; starch, ko00500; terpenoids, ko00900; tetrapyrrole, ko00860 (Table S1). SUF (sulfur mobilization) is not present inKEGG. Tat; Twin-Arginine Translocation pathway, Tic/Toc; Translocon at the Inner/Outer envelope membrane of Chloroplasts.doi:10.1371/journal.pgen.1004355.g004



modification, and protein turnover. Most surprisingly, Helicospor-

idium has not seen an increase in the complexity of transporter

families, which might be expected of a parasite as host dependence

grows; instead, this functional class is most reduced in Helicospor-

idium compared with Coccomyxa and Chlorella.

Gene family expansions: ChitinasesTaking an opposing view, to what gene families may have

expanded due to the adaptation to parasitism, revealed a single

obvious expansion of functional significance. A total of 14 genes

with putative glycosyl hydrolase (GH) activity were identified

throughout the Helicosporidium genome and were also found within

its transcriptome (Table 2). All of these proteins appear to belong

to the GH18 chitinase family, whereas plant chitinases normally

come from the GH19 family. The Chlorella genome encodes two

GH18 and one GH19 chitinase, and these are assumed to be

involved in the remodelling of its cell wall, which has been

experimentally demonstrated to contain chitin [17,26]. The GH19

chitinase in Chlorella was acquired by horizontal gene transfer from

a large DNA virus [17] and is not found in either the

Helicosporidium or Coccomyxa genomes. Conversely, the Chlorella

GH18 chitinases are found across the green algae. In Helicospor-

idium, the extra 12 copies appear to have been generated by recent

duplication events. We found no evidence for the transfer of

genetic material from insects in the Helicosporidium genome or

transcriptome. A total of 13 of the 14 Helicosporidium chitinases

contain the Dx2DxDxE/P motif essential for chitinolytic activity.

Insect and bacterial chitinases with experimentally confirmed

catalytic activity also contain one or more of three additional

motifs: Kx6GG, MxYDx(x)G, and Gx3Wx2DxD [27–29]. Thir-

teen of the chitinases in Helicosporidium contain one or two of these

additional motifs and demonstrate high conservation in their

Figure 5. Evolutionary gene network analysis showing functional contractions in Helicosporidium relative to Chlorella and Coccomyxa.The connected components depicted here represent a small portion of the genes involved in each of these functional categories. Helicosporidium,Coccomyxa and Chlorella are represented by orange, blue and green nodes, respectively. Interactions between the different components are denotedby gray lines connecting the nodes.doi:10.1371/journal.pgen.1004355.g005



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orientation (Table 2, Figure S3). However, in addition to

H632_c1867p1 which appears to lack this domain, three show

substitutions within the Dx2DxDxE motif and may be therefore be

inactive.

Another class of proteins that might be expected to be relevant

to the origin of parasitism in Helicosporidia are, unfortunately, the

unidentified or ‘unique’ ORFs. To see whether these represented a

high proportion of the genes in Helicosporidium, we identified

predicted proteins of at least 100 amino acids that are not found in

Coccomyxa and Chlorella, which resulted in 882 distinct proteins

(Data S12). A small number of these proteins have clear or

putative homologs in the Volvox and/or Chlamydomonas genomes

but the vast majority are unique and have no known homologs.

The few cases with homologues in Volvox and Chlamydomonas are

sulfotransferases, glycosyltransferases or hydrolases with chitinase

activity (as mentioned above), a 2-oxoglutarate Fe(II)-dependent

oxygenase, and a cyclin. In contrast, the predicted proteins

without known green algal homologs could not be assigned to any

putative function in PFAM homology searches at an E-value cut-

off of 1e-10. Five of these unknown proteins (H632_c233p3,

H632_c338p0, H632_c531p0, H632_c1976p1, H632_c4072p0)

display mid to low similarity with bacterial sequences of unknown

function, but they are unambiguously encoded on contigs

encoding clearly eukaryotic genes, and are therefore not bacterial

contaminants. From the transcriptome, we can also conclude that

the majority of these proteins are expressed, with 585 of the 882

found in transcriptome data at an E-value threshold of 1e-15.

Horizontal gene transfer from/to virusesWe identified two transcripts sharing a high identity with viral

sequences (E-value threshold of 1e-40), the closest relatives being

Paramecium bursaria Chlorella viruses (PBCV) and Acanthocystis turfacea

Chlorella viruses (ATCV). These two transcripts were also identified

in the Helicosporidium genomic contigs, and both are assembled with

Helicosporidium nuclear genes and are therefore not the result of

viral contamination. The first transcript (a374428r16) codes for a

dUDP-D-glucose 4,6 dehydratase that is also found in the

genomes of Chlorella, Coccomyxa and various other green algae,

and has been reported as an example of host to virus horizontal

gene transfer (HGT) [30,31]. The second transcript (a28443r121)

encodes a D-lactate-dehydrogenase and is also found across the

green lineage. Phylogenetic analyses including the closest D-

lactate-dehydrogenase sequences clusters the green algal sequences

with homologues from nucleocytoplasmic large DNA viruses that

infect them, with strong bootstrap support (Figure 6). Overall, this

suggests D-lactate-dehydrogenase represents another case of host-

virus HGT in the green algal lineage.

Discussion

A lack of loss in the evolution of parasitismA recurring theme in the evolution of parasite genomes and

parasitism in general is reduction. In parallel with the more

constructive process of developing sometimes sophisticated mech-

anisms to invade and take advantage of their hosts, parasite

evolution generally involves the selective pruning of biological

functions that are no-longer mandatory to survival in the host.

This reduction has occurred independently many times during the

evolution of parasites, and is generally reflected in their genomes,

streamlined sometimes solely by a loss of genes, but other times

also by an overall shortening of coding and intergenic regions as

well as a loss of introns, resulting in much smaller and compact

genomes than those of their free-living relatives [9]. In very few

cases have photosynthetic algae made this transition, but the

famous exception is the apicomplexans, including the malaria

parasite Plasmodium. Here the same process has taken place and

although the plastid has been retained, it has been reduced to a

cryptic form that lacks most of its ancestral metabolic pathways

and has retained nothing whatsoever related to carbon metabo-

lism.

Helicosporidium breaks from these trends in significant ways. With

an almost perfect conservation of the core green algal metabolic

pathways, its genome is small, but can hardly be considered

reduced, which may reflect its relatively recent adaptation to

parasitism. Particularly surprising is the retention of an almost

complete pathway for carbon fixation. Helicosporidium has lost

nearly all genes associated with light harvesting, photosystems, and

chlorophyll biogenesis, so how it uses carbon fixation pathways is

an interesting question. Carbohydrate storage, and in particular

the use of starch could be the main driver behind the retention of

carbon fixation genes. Successful parasites often sequester

resources from their host [32] and converting simple sugar

molecules to large starchy polymers polarizes the directionality of

carbohydrate exchanges. Exactly how efficient are the Helicospor-

idium permeases at sugar uptake and other energy-related

metabolites from the immediate environment is unclear; consid-

ering that this is one of the gene families of reduced complexity,

but such sequestration may be a prerequisite for survival.

The coding density of the Helicosporidium genome is only

marginally higher than its closest relatives, and almost on par

with those of a number of free-living prasinophytes (Table 1;

Figure 1). Green algal genome sizes range widely across lineages

(Figure S1), and expansions as well as contractions are likely to

have occurred several times independently. Accordingly, we

cannot currently distinguish between a reduction of DNA

packaging/replication/translation pathways in Helicosporidium, an

expansion of these pathways in Chlorella/Coccomyxa, or a combi-

nation of both. A better phylogenetic framework as well a greater

sampling depth of green algal nuclear genomes will be required to

polarize the directionality of this event, but the reduced gene

family complexity observed in picoprasinophytes [33–35] does

argue in favour of an expansion from a lean ancestral state.

Despite its small size, the Helicosporidium genome does feature

one small expansion. Chitinases are uniformly rare in green

algae where genomic data are available, suggesting this gene

family is ancestrally limited in the green algae (which fits with

its likely function in remodelling a minor component of the cell

wall in some species). The expansion of chitinases in

Helicosporidium over its sister taxa Chlorella and Coccomyxa

therefore likely represents a unique adaptation that directly

resulted from or even contributed to its parasitic lifestyle in

insect hosts. The Helicosporidium infection mechanism consists

of the oral uptake of cysts, dehiscence in the midgut lumen,

ingression through the midgut, and entry into the hemocoel

where the organism multiplies. In a number of insect parasites,

including the malarial parasite Plasmodium, the presence of

exogenous chitinases from the GH18 family within the

arthropod midgut tract is either mandatory for infection or

associated with increased pathogenicity by allowing pathogens

to pass through the peritrophic membrane [36,37]. Disruption

of chitinase activity with blocking agents interferes with the

sporogonic development of malarial parasites, which is

restored upon addition of exogenous chitinases [38]. One

may speculate that the Helicosporidium chitinases serve multi-

functional roles and operate at various stages in vivo.

Potentially, chitinases sequestered in the cysts are activated

by insect digestive proteases, and then digest the cyst wall to

initiate the release of the invasive filament cell in the gut



lumen. Alternatively, the chitinases released by the ovoid cells,

may loosen the chitin matrix of the peritrophic membrane,

allowing the ingress of the filamentous cell into the ectoperitrophic

space. It is important to note that members of the GH18, in addition

to binding to and digesting the chitin can also target various

GlcNAc-containing glycans that comprise various exocellular

matrices including insect basement membranes. Binding to such

substrates may aid in the establishment of infection on the hemocoel

associated tissues. Finally the production of these enzymes could

soften the exoskeleton, which could play a role in the egress of the

infectious cyst stage from diseased insects.

Materials and Methods

Tissue culture and DNA/RNA purificationVegetative cells of the Helicosporidium sp. (ATCC50920) a

parasite of the black fly Simulium jonesi [1,15] was propagated in

stationary cultures of sabouraud dextrose broth for five days at

27uC. Cells were harvested by centrifugation (5,000 rcf for

10 min) and pellets resuspended in a minimal volume sterile

H2O and used for nucleic acid extraction. For genomic DNA

preparation, a total of 26109 cells were suspended in the yeast

lysis buffer (Epicentre Biotechnologies, Madison WI) and

homogenized with Bead Beater technology. The extracted

nucleic acid phase was treated with DNase free RNase and

subjected to chloroform phenol extraction, precipitated with

ethanol, and suspended in TE buffer. A total of 14.1 mg high

molecular weight DNA were recovered and submitted for

sequencing. For total RNA extraction the resuspended harvested

cells were immediately added to liquid N2 and ground with mortar

and pestle to break the outer pellicles. The resulting frozen cell

powders were processed initially with TRizol Reagent then

processed with Purelink RNA Mini kit (Ambion). Column eluants

were treated with RNase-free DNase and analysed with the 2100

Bioanalyzer (Agilent Technologies, Inc). Samples (10 mg) producing

RIN values of 8.9 to 9.2 were selected for subsequent sequence

analysis (see below).

DNA and RNA sequencingTotal DNA and RNA from Helicosporidium sp. ATCC 50920

(mitochondrial, plastid and nuclear) were sequenced by

Fasteris SA (Plan-les-Ouates, Switzerland) using the Illumina

platform. Two independent DNA sequencing runs were

performed. In the first, 18,618,066 reads (54-bp paired ends;

323-bp inserts; average standard deviation, 19) totaling

1,005,375,564 bases were sequenced with the Illumina GA-

IIx platform and the Chrysalis 36 cycles v 4.0 sequencing kit.

In the second, 17,110,904 reads (51-bp paired ends; 241-bp

inserts; average standard deviation, 64) totaling 872,656,104

bases were sequenced with the Illumina HiSeq 2000 platform

and the TruSeq chemistry. Total RNA was sequenced using

the Illumina directional mRNA-SEQ protocol with the

Figure 6. Maximum likelihood tree of the D-lactate dehydrogenase transferred by HGT between green algae and their viruses. Thebest protein ML tree is shown here. Bootstrap support is indicated above the corresponding nodes. Fungal and bacterial tips were converted totriangles for concision. ATCV, Acanthocystis turfacea Chlorella viruses; PBCV, Paramecium bursaria Chlorella viruses.doi:10.1371/journal.pgen.1004355.g006



Illumina HiSeq 2000 platform and the TruSeq chemistry. A

total of 83,075,963 reads (100-bp single ends) were generated

(8,307,596,300 bases total). Read quality for each Illumina

data set was assessed with FastQC (version 0.10.1;

Babraham Bioinformatics, Babraham Institute [http://www.

bioinformatics.babraham.ac.uk]).

Genome assemblyPaired-end reads were assembled de novo with Ray [39] 2.0.0

rc8 using iterative k-mer values of 21 to 31 on 8 processing

cores (2 Intel Xeon E5506 CPUs at 2.13 GHz) with a

maximum RAM allowance of 96 Gb. The resulting contigs

were filtered by size with sort_contigs.pl (Advanced Center for

Genome Technology, University of Oklahoma [www.genome.

ou.edu/informatics.html]), and contigs shorter than 500 bp

were discarded. The contigs of at least 500 bp were conserved

for downstream analyses. The 500+-bp contigs were used as

canvas to generate a BLAST [40] database with MAKE-

BLASTDB from the NCBI BLAST 2.2.26 package, the

mitochondrial and plastid contigs were identified by BLAST

homology searches using the mitochondrial (GenBank acces-

sion number NC_017841, [41]) and plastid (GenBank acces-

sion number NC_008100, [20]) genomes as queries, and

separated from the nuclear contigs. Putative contaminants

were assessed by homology searches against the NCBI non-

redundant database.

Transcriptome assemblyRNA-Seq reads were filtered using a sliding-window quality

approach with Sickle (Bioinformatics Core, University of

California, Davis [https://github.com/najoshi/sickle]) under

the default parameters, and the overall read quality reassessed

after filtering with FastQC. Illumina adapter sequences were

then removed from the filtered sequences using custom Perl

scripts, and PolyA-tails were removed from the reads with

TrimEST from the EMBOSS [42] 6.4.0 package. The filtered

transcriptome reads were assembled with Trinity’s Inchworm

module with a maximum RAM allowance of 90 Gb (–JM 90G)

on 8 processing cores (2 Intel Xeon E5506 CPUs at 2.13 GHz).

Contigs were filtered by size with sort_contigs.pl and contigs of

at least 250 bp were selected for downstream analyses.

Transcriptomic contigs were mapped on the genomic ones

with GMAP version 2014-01-21 [43] using the default

parameters.

Genome annotationThe nuclear contigs of at least 500-bp in length were sorted by

size and renumbered incrementally using customs Perl scripts.

Contigs were then processed with the Maker 2.11 annotation

gauntlet [44,45] using the Chlorella gene model as implemented in

Augustus 2.5.5 [46]. The resulting GFF annotations files were

processed, curated, and converted to GenBank annotations files

using custom Perl scripts. Putative functions were assigned using

homology searches against the PFAM database (E-value threshold

of 1E-30; Table S5). Transposable elements were searched for

with RepeatMasker [http://repeatmasker.org] using Repbase

version 20130422 [47].

Genome size estimationIllumina reads from the mitochondrial and plastid genome were

first filtered out from the total dataset with bowtie 0.12.9 [48] using –

un and –al the flags against indexes built from the organelle sequences.

Filtered nuclear reads were then mapped with bowtie against the

5,666 contigs ($500 bp) with the –S flag, and the coverage estimated

from the SAM file with Tablet 1.12.12.05 [49] and the coveragestat.py

python script. The genome size was then estimated using the following

formula: [# of reads X read length]/coverage.

Pathways mapping and network analysesKEGG metabolic pathway maps for the green algae

Chlamydomonas reinhardtii, Volvox carteri, Ostreococcus tauri and

Ostreococcus lucimarinus were retrieved from the KEGG pathway

databases [50,51], the proteins sorted accordingly, and then

used as queries for homology searches against the Helicospor-

idium, Chlorella and Coccomyxa proteomic, genomic and tran-

scriptomic datasets (the Chlorella and Coccomyxa data was

retrieved from the JGI website). BLASTP and TBLASTN

searches were performed using E-value thresholds of 1E-10

and 1E-05, respectively. Genes not found in searches against

any of the three datasets were considered absent from the

corresponding organism. Network analyses were performed

according to [25]. Specifically, all possible edges were drawn

between pairs of genes if their reciprocal BLASTP compari-

sons to one another met all of the following conditions: E-

value,1E-10, minimal hit identity .20, at least 20% of the

shortest gene’s length had identical residues in the match, and

the hit length .20 amino acids. The network was then filtered

to include underrepresented Helicosporidium genes compared to

Coccomyxa and Chlorella. Functional annotations for the genes

comprising each connected component (GenBank, KOG,

KEGG, Interpro [52], and Pfam) were used to characterize

each connected component by its inferred biological function.

Plastid-targeted proteins from GreenCut2’s Table S2 [19] were

extracted from the corresponding Chlamydomonas reinhardtii

(version 3.1) and Arabidopsis thaliana (http://www.arabidopsis.

org/) protein catalogs and converted to custom BLAST

databases with MAKEBLASTDB from the NCBI BLAST

package. Helicosporidium, Chlorella and Coccomyxa were searched

independently against both GreenCut2 databases with

BLASTP (proteins) and TBLASTN (genome and transcrip-

tome) using E-value thresholds of 1E-10 and 1E-05, respec-

tively.

Chitinase identificationPutative glycosyl hydrolases identified in the Helicosporidium

genome were annotated for catalytic and chitin binding domains

using SMART 7 [53] and endo-proteolytic sites often located

within developmental insect chitinases were identified with

ePESTfind [54]. The glycosyl hydrolase catalytic domains were

annotated manually for the presence and orientation of key amino

acid motifs. Secretory signal motifs were searched for with TargetP

1.1 [55] and PredAlgo [56].

Phylogenetic analysesAmino acid sequences retrieved from GenBank were aligned

with the L-INS-I algorithm from MAFFT 7.029b [57]. Phyloge-

netic models were selected with ProtTest 3.2 [58]. Maximum

Likelihood phylogenetic reconstructions were performed with

PHYML 3.0 [59] under the LG+C4+I model of amino acid

substitution [60].

Data depositionThe Helicosporidium data was deposited at DDBJ/EMBL/

GenBank under NCBI BioProject ID PRJNA188927 and

accession AYPS00000000. The version described in this paper

is version AYPS01000000. The predicted proteins and RNAs



http://www.bioinformatics.babraham.ac.uk

http://www.bioinformatics.babraham.ac.uk

www.genome.ou.edu/informatics.html

www.genome.ou.edu/informatics.html

https://github.com/najoshi/sickle

http://repeatmasker.org

http://www.arabidopsis.org/

http://www.arabidopsis.org/

are also available in Data S13 and S14, respectively. All

custom Perl scripts are available on GitHub (https://github.

com/JFP-Laboratory).

Supporting Information

Data S1 Transposable elements found in the Helicosporidium

contigs. Repeats were searched for with RepeatMasker [http://

repeatmasker.org] using Repbase version 20130422.

(TXT)

Data S2 PFAM annotations of the Figure 2 proteins. The

searches were performed with an E-value cut-off of 1.0; Figure 2

proteins that are absent from this file are hypothetical proteins.

(TXT)

Data S3 Orthology map of the valine/leucine/isoleucine

biosynthesis pathway (ko00290) retrieved from KEGG [50,51].

Genes that are present in Helicosporidium are indicated by yellow

boxes. Genes that are absent from Helicosporidium but present in

Chlamydomonas are indicated by red boxes. Genes that are absent

from both Helicosporidium and Chlamydomonas are indicated by

empty boxes.

(PNG)

Data S4 Orthology map of the phenylalanine/tyrosine/trypto-

phan biosynthesis pathway (ko00400) retrieved from KEGG

[50,51]. Genes that are present in Helicosporidium are indicated

by yellow boxes. Genes that are absent from Helicosporidium but

present in Chlamydomonas are indicated by red boxes. Genes that

are absent from both Helicosporidium and Chlamydomonas are

indicated by empty boxes.

(PNG)

Data S5 Orthology map of the starch and sucrose metabolism

pathway (ko00500) retrieved from KEGG [50,51]. Genes that are

present in Helicosporidium are indicated by yellow boxes. Genes that

are absent from Helicosporidium but present in Chlamydomonas are

indicated by red boxes. Genes that are absent from both

Helicosporidium and Chlamydomonas are indicated by empty boxes.

(PNG)

Data S6 Orthology map of the carbon fixation pathway in

photosynthetic organisms (ko00710) retrieved from KEGG

[50,51]. Genes that are present in Helicosporidium are indicated

by yellow boxes. Genes that are absent from Helicosporidium but

present in Chlamydomonas are indicated by red boxes. Genes that

are absent from both Helicosporidium and Chlamydomonas are

indicated by empty boxes.

(PNG)

Data S7 Orthology map of the porphyrin and chlorophyll

metabolism pathway (ko00860) retrieved from KEGG [50,51].

Genes that are present in Helicosporidium are indicated by yellow

boxes. Genes that are absent from Helicosporidium but present in

Chlamydomonas are indicated by red boxes. Genes that are absent

from both Helicosporidium and Chlamydomonas are indicated by

empty boxes.

(PNG)

Data S8 Orthology map of the terpenoid backbone biosynthesis

pathway (ko00900) retrieved from KEGG [50,51]. Genes that are

present in Helicosporidium are indicated by yellow boxes. Genes that

are absent from Helicosporidium but present in Chlamydomonas are



(PNG)

Data S9 Orthology map of the carotenoid biosynthesis pathway

(ko00906) retrieved from KEGG [50,51]. Genes that are present

in Helicosporidium are indicated by yellow boxes. Genes that are

absent from Helicosporidium but present in Chlamydomonas are



(PNG)

Data S10 Orthology map of the unsaturated fatty acids

biosynthesis pathway (ko01040) retrieved from KEGG [50,51]. Genes

that are present in Helicosporidium are indicated by yellow boxes. Genes

that are absent from Helicosporidium but present in Chlamydomonas are

indicated by red boxes. Genes that are absent from both Helicosporidium

and Chlamydomonas are indicated by empty boxes.

(PNG)

Data S11 Orthology map of the protein export pathways

(ko03060) retrieved from KEGG [50,51]. Genes that are present

in Helicosporidium are indicated by yellow boxes. Genes that are

absent from Helicosporidium but present in Chlamydomonas are



(PNG)

Data S12 Helicosporidium proteins that are not found in other

trebouxiophytes. Predicted proteins of at least 100 amino acids

that are not found in the trebouxiophytes Chlorella and Coccomyxa

[E-value threshold 1E-05] are included in this file. The vast

majority of these proteins are unique and have no known

homologs.

(TXT)

Data S13 Predicted proteins in Helicosporidium. The proteins

were predicted from the genomic contigs with MAKER 2.11 using

the Chlorella gene model as implemented in Augustus 2.5.5.

(TXT)

Data S14 Helicosporidium RNA-Seq contigs. The Helicosporidium

RNA-Seq data was assembled with Trinity’s Inchworm module

from quality filtered reads.

(FASTA)

Figure S1 Green algal genome sizes in the phylum Chlorophyta.

Sequenced green algae are labelled in black with the source

indicated between parentheses. Estimated values based on nuclear

DNA content from Kapraun [61,62] are color-coded according to

their respective group. Blue, U, Ulvophyceae; Orange, C,

Chlorophyceae; Green, T, Trebouxiophyceae; Purple, P, Prasino-

phyceae. Lower and upper estimates, when present, are shown in

dark and light colors, respectively. Note that the real and estimated

sizes of the Ostreococcus tauri genome differ by an order of magnitude.

(PDF)

Figure S2 Heme and chlorophyll pathways in Helicosporidium.

Genes present in Helicosporidium are indicated in orange. Genes

absent are shown in red. This simplified schema is derived from

KEGG pathway KO00860 [50,51].

(PDF)

Figure S3 CLUSTALW alignment of the Helicosporidium chit-

inase Gly18 catalytic domains. Conserved motifs are shown in

blue. BLOSUM, Gap open penalty: 35, Gap extend penalty 0.75.

(TIFF)

Table S1 Distribution of the Chlamydomonas reinhardtii KEGG

metabolic pathways in the trebouxiophytes Helicosporidium, Chlorella

and Coccomyxa. The Chlamydomonas, Chlorella and Coccomyxa proteins

and genomic/transcriptomic contigs are labelled according to the



https://github.com/JFP-Laboratory

https://github.com/JFP-Laboratory



official headers provided in the JGI data (http://genome.jgi.doe.

gov/).

(XLSX)

Table S2 Enzymes that are bypassed or lost in Helicosporidium

non-photosynthetic pathways. The Dicer [K11592; EC:3.1.26.-]

and Argonaute [K11596] proteins absent from Helicosporidium are

not included in the table; the only KEGG pathway currently

including Dicer/Argonaute proteins relates to cancer.

(XLSX)

Table S3 Underrepresented Helicosporidium metabolic pathways

in evolutionary gene network analyses. The connected compo-

nents represent a small portion of the genes involved in each of

these functional categories.

(XLSX)

Table S4 InterProScan5 analyses of the connected components

from Figure 5. Helicosporidium; sheet 1, Chlorella; sheet 2, Coccomyxa; sheet

3. The complete InterProScan5 format definition can be found at

[https://code.google.com/p/interproscan/wiki/OutputFormats].

(XLSX)

Table S5 GenBank-annotated proteins in Helicosporidium sp. Only

proteins displaying matches at an E-value cutoff of 1E-30 against

PFAM were annotated for release in the GenBank database.

(XLSX)

Author Contributions

Conceived and designed the experiments: JFP DB PJK. Performed the

experiments: JFP NAB DB. Analyzed the data: JFP NAB. Contributed

reagents/materials/analysis tools: JFP DB CL. Wrote the paper: JFP NAB

DB PJK.

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